Collecting insects

This is a little article I wrote to help novice entomologists understand the basic principles behind making an insect collection.

Why do we collect insects?

It’s a fair question – especially when there are such amazing cameras these days and forums all over the web where experts will seemingly put a name to just about anything. However, photographs of living insects very often do not show the parts that confirm the species, like the genitalia or minute bristles and surface sculpturing. It is possible to identify some insects from photos but even a lot of common insects might only be identifiable to genus or the expert might say “probably X” or “possibly Y” – so they have no way to be really sure.

But do we need to know exactly what something is?Why can’t we leave wildlife to get on with it? Well, investigating the world around us is how science has brought civilisation to the level that we have now and the basis of all sciences is to be able to identify what you are looking at. Imagine how much the science of chemistry would have advanced if chemists couldn’t tell gold from lead!

If we can identify species and understand how their populations are being changed by human interaction (eg. global warming) then we have the best chance to help them in the long term. We can monitor their populations and distributions and better target conservation efforts to the exact causes of decline.

Can insect populations survive collection? What if we kill the last one? Collectors these days very rarely take more than a few individuals of each species because they are just sampling a particular area or taking reference material. The days of the Victorian-style collector, taking long series of rare species to fill gaps in his collection, are long gone.

The taking of a few specimens in any location is not going to make any difference to a population of invertebrates because they exist in such large numbers. Insects reach maturity quickly and lay lots of eggs and so their capacity to recover even large population disasters (like a fire or flood) is far greater than with vertebrates, which breed far slower and exist in smaller populations. Predators like birds and other insects will eat many hundreds (even thousands) of insects in a day and this is all considered normal so the effects of a few entomologists really doesn’t amount to anything.

Stuart Ball & Roger Morris once did a highly scientific “mark & recapture” experiment to ascertain population density and movements within a particular habitat. Their results showed that no matter how hard they caught their flies (hundreds were captured) and marked them, the fact that they re-caught so few marked ones showed that they were only affecting a mere 16% of the population. This shows how tiny would be the effect of removing a few individual specimens.

Isn’t killing things immoral? This is a difficult and highly subjective question to which you can find many answers. I have never met an entomologists who enjoy the act of killing but we are scientists who understand that the effect of our science is likely to be far more beneficial in the long term because the knowledge gained will go to help the species we are collecting and the habitats they live in. When entomologists euthanize specimens they do so in the most humane ways and always prior to pinning so that there is no actual cruelty involved.

Furthermore, if one is so worried about the avoidable death of invertebrates then consider the effect that we all have on the world around us. We kill hundreds of insects in our cars and I need not even mention the use of insecticides in the manufacture of the food we all eat. Indirectly we also contribute to the death of millions with global warming, intensive agriculture and mining. There are few ways that we don’t affect invertebrates but if we are careful and we try to mitigate this wholesale destruction then insects will thrive.

Basic reasons that you might want to collect

Entomologists frequently have to catch and kill a few insects to identify them because it isn’t possible to key some orders without being able to examine them in minute detail under a microscope. Accurate taxonomy & identifications / records are a vital cornerstone of entomological, environmental & conservation sciences because they provide vital distribution and biodiversity information. Taking regular specimens over time will show you variations in the population and without the work of the old collectors we wouldn’t have been able to detect the changes that had happened to species like the Peppered Moth.

Once a specimen has been identified it is almost always kept for future reference in a personal reference collection – they are not thrown away. This is so that the record can be backed up by physical evidence that can be examined later by other workers and it can also be very useful to have reference material that you can compare to any newly identified specimens. This is especially true when you are starting to learn a new group and are not familiar with the range of the fauna. Having specimens also allows us to revisit the work that we did years previously when we were young and inexperienced. We all make mistakes and being able to show the specimens to someone else (or your better self) allows corrections to be made.

Things to think about

When collecting for science (as opposed to ‘pretty displays to hang on the wall’) there are only usually 2 things to think about:

How can I protect the specimen in the long term?

How can I present the specimen so that it can be examined and identified as easily as possible?

With each group of insects you will find different answers to these questions, depending on how hard/soft the specimen is and which features are used in the identification keys. For instance, some keys assume that specimens are preserved in alcohol (eg. spiders / molluscs) and others assume they will be dry (eg. most insects).

In the following article I will assume that we are talking about Diptera & Hymenoptera (my specialities). Also, please remember that a lot of this is my personal preference – other people might/will disagree!!

Obtaining specimens

You can start with dead insects you find at home or in the countryside but they will usually be too dry & brittle to pin immediately. Place them in a plastic box on wet tissue paper for 24 hours to make the soft enough.

If the specimens come from wet traps – traps that collect into liquid (eg. malaise or pitfall traps) then the insects will usually to be supple enough to pin immediately.

If you collect live insects then the easiest and most humane way to bump them off is to put them in a domestic freezer (-18c and below) for a few hours. Then when you take them out let them defrost for an hour before pinning.

Mounting specimens

I side-pin and stage-mount most small to medium-sized specimens because it has many clear benefits for Diptera & even some Hymenoptera. The micro-pin goes into the side of the fly at a slight angle so that it doesn’t damage the same feature on both sides – all manipulated by fine-pointed forceps. The fly is then pinned to a block of foam and the legs, head & wings all arranged to show off the essential features. If the specimen is male you can very easily hook-out the genitalia and use more micro pins to hold the bits out while the specimen dries.

When the specimen is dry I make a ‘stage’ using a thin strip of plastazote (high density foam) and push a 38mm, continental-sized, entomological pin (size 3 or above) through one end so that the stage sticks out at 90-degrees. The micro pin with the specimen is then pushed firmly into the foam stage. The stage is positioned half way up the stage-pin, to allow room underneath for labels, but not too close to the top of the pin where chubby fingers are likely to brush against the specimen. (see right).

Staging combined with side-pinning protects small/medium specimens very well while displaying as many features as possible. The stage-pin is strong and easy to hold/manipulate when moving the specimen and the stage absorbs most vibrations.

Alternatively you can direct-pin (see right), either laterally or dorsally (‘top-down’) but I only do this with very large specimens (eg. Tachina spp. or bigger) where you can use a relatively thick pin (such as a #1 or #2 thickness). Never use the finest thickness (0 and 00) of 38mm pins because they are almost impossible to push into older cork boxes and also bend and twist when you push them into foam, which can cause the specimen to crack or twang and loose legs and antennae.

Another popular technique (especially with hymenopterists) is gluing the insect to strong card but I feel that very fine micro pins are:

much easier to use (glue is very messy)

allow for moving later (glue can be dissolved but it isn’t easy)

need not destroy much/any surface features

Also, I like to have a single mounting method for most of my specimens and glue doesn’t work well for groups like Diptera, which have fragile bristles.

Knock-on benefits of staging

Coincidentally, micro-pinning lends itself very well to bulk-collecting of groups that take a while to identify.

I usually go out for a day in the field and catch maybe 50 insects of various orders and families. I also work with Malaise traps, which can generate vast numbers of specimens on a single day of sorting. This is just too many to work with in a single day so I reduce the work I have to do by creating batches of insects.

I simply pin all the specimens with micro pins and put them in a flat, clear, plastic box with a sheet of foam at the bottom (see right) with a data label containing all the collection data for that group of insects. The specimens stay here to dry and allow me to continue working on other batches in the meantime.

When I have time to identify them I pin a specimen to a new stage; identify it; and put the specimen’s labels (copy of the data & a separate label for the name) on the big stage-pin, under the stage. The specimen is then ready to be transferred to the collection-proper. The other specimens may stay in the plastic box for years waiting for me to have time to work on them but while they are in this state they take up very little room and they are very well protected from dust and damage.

This all fits neatly into a plastic ‘tupperware’ box for storage and transportation.

Pins

I use a variety of different pins for different jobs:

Continental length (38mm) pins for use as stage-pins (in size 3 – 5 – approximately .55mm diameter) or sometimes for direct-pinning very large specimens (in size 1 or 2). If the specimen is small enough to need a thinner pin then you should be staging it.

Micro pins for direct-pinning most of my specimens:

D3 size (0.01″ x 10mm) for larger specimens

A1 size (0.0056″ x 10mm) for anything small

Microscope & light source

A good binocular microscope with lighting system is an essential part of any entomologist’s equipment and is probably the most expensive thing you will buy. Ideally you want to invest in something that has very good optics and a good, bright white light source. You don’t need to have very strong magnification – about 5x-40x is perfectly adequate for most flies & wasps. When you are ready to buy a microscope the main thing is to try out plenty of different models and test them with a few of your own specimens.

I currently use 2 setups:

Leica S8APO with chinese 244-LED ring-light – a lovely microscope but a bit expensive for most beginners to buy (~£4000 new from a Leica dealer). Fitted with 10x wide-field eyepieces the zoom range is 10x to 80x and the camera port on the top coupled with the fine-focus rack-stand allows you to take stacked photos of whatever you are looking at. The optical quality is absolutely superb.

My first microscope was a MEIJI EMZ zoom binocular microscope with a zoom range of 0.7x – 4.5x and 10x eyepieces giving me 7x – 45x total magnification. The light for this scope is a small Mini-Fluor fluorescent light attached to the microscope itself. I find it very good and easy to work with and the most important thing – no eye-strain after working with it for hours on end!

The 244-LED ring-lights can be used on virtually any microscope and can be bought on eBay for around £50 – just shop around.

Accessories

I like to have one of the eyepieces fitted with a graticule – a grid or ruled line that helps you measure relative lengths of objects on the specimen. They fit inside one of the eyepieces and are fiddly to put in so the standard technique is to either have it in a spare eyepiece or just leave it in one of your main eyepieces all the time.

Secondary lighting can also be very useful when peering into dark corners of a small specimen. An easy and cheap solution to this is to get a small, household angle-poise desk lamp fitted with an energy-saving fluorescent or LED bulb. These can be pointed to shine directly at the specimen from any angle and the tube is never usually hot so you don’t risk burning the back of your hand – or melting the specimen’s wings!!

Working with genitalia

Some identifications rely on being able to examine the male genitalia and you can usually use the genitalia as a good confirmatory character. For this reason I always recommend that if you have a male you should always try to hook out the genital capsule and aedeagus when the specimen is fresh. You never know when they will be important but it’s much easier to do while they are soft and pliant, than try to relax an old specimen – a little bit of work early-on will save you a great deal of hassle later.

All you need to do is to use a fine micro-pin and gently tease open the genital capsule. You should find that it hinges on its dorsum and you can hold it open with a few crossed micro-pins until the specimen has dried and is set (see right). Sarcophagids are an extreme example because they have such huge genital capsules but they are also a good example because the work is incredibly hard to do unless you open it while the specimen is still flexible.

For some groups (eg. sarcophagids) it helps if you can also locate and tease out the aedeagus, a small chitinous filament that’s analagous the penis in higher animals. This is pretty small and of course it is the most internal & anterior part of the external genitalia so it is difficult to get to – but it isn’t impossible and it makes the genitalia preparation complete. It usually pops out if you extend the cerci fully – about 90-degrees to the body.

Sometimes you will have to remove the genitalia completely and in these cases they can be clarified in a weak solution of potasium hydroxide (KOH) but it isn’t usually necessary. Be very careful with this chemical – it is strongly caustic and care should be taken to keep it away from the skin and eyes!

I normally store separated genitalia in little plastic capsules that can be pinned beneath the specimen, just above the data label (see right). The genitalia are suspended in a solution of glycerine. This system protects them but also allows them to be examined later – though it can be tricky to see much through plastic and glycerine so you have be prepared to remove them.

Alternatively, some people choose to store the genitalia in a water-soluable, transparent resin DMHF. When dry DMHF becomes a hard, long-lasting and transparent droplet that both protects and holds the genitalia, yet allows it to be examined and can be dissolved later if necessary.

Labels

All specimens should have at least 2 labels:

the data label (first under the stage) containing:

Place of capture (region, place name & map reference)

Date of capture (often with the month in roman numerals to prevent confusion between American and European date formats)

If the specimen was reared from a host or collected as a larvae you might also want to give it a rearing label (under the data label) containing:

host name

date of capture

any other notes relating to the rearing

These labels are made from card or thick 160gsm paper so that they don’t drop or rotate on the pin. Never use thinner paper – it won’t grip the pin properly and it will start to spin round and need replacing very quickly.

I normally print as much of the static information as I can, using laser or inkjet printers fitted with indellible inks. At the begining of each year I print:

A sheet of part-completed data labels for each of my favourite collecting sites containing all the information except the day & month of collection.

Sheets of completed det labels for the groups I study most frequently. I collect a lot of tachinids and I am also asked to det tachinids throughout the year so it makes sense for me to carry around sheets of pre-printed det labels to save me having it write them each time. Obviously, I print more labels for the common species and less of the rarities.

A sheet of ‘blank’ det labels (with just my name and the current year) which I will use for all non-tachinids.

Resist the urge to write additional information on the back of the labels – nobody will see it or check it and the specimen will have to be picked up to read it (increasing the potential for handling damage). If you have additional information then print a bigger label and print it on thicker card – the larger labels will actually protect your specimen better!

Storage

As long as insect specimens are kept dry and pest free they will last for centuries so the storage requirements can be very simple indeed. You could just start with sturdy, air-tight tupperware/food storage boxes and glue some 6mm foam onto the bottom to hold your pins. You could arrange them so that each box holds 1 genus or just mix specimens together – whatever you can cope with without loosing specimens in a muddle!

Most amateurs use purpose-built, wooden store boxes (see right). Second-hand boxes can be bought from most natural history museums (from around £10 – contact Max Barclay at the Natural History Museum in London) and you can just line the bottom with a 6mm or 9mm sheet of Alveolit foam. You can also buy new boxes from entomological dealers but they can be a little expensive.

I used store boxes for many years but as my reference collection started to reach 300 species with 2000 specimens it became too difficult to manage. Inserting new species often meant that I had to move lots of specimens around and that brought damage and took a long time. Also, store boxes are not very air-tight and so pests can get in and if not noticed for a while can cause immense damage.

Drawers & unit-trays

Once your collection spans 5 or more store boxes it starts to become very unweildy to curate and refer to. Each time you insert a new species you have to shuffle the rest up or down in the box to make space and each time you do this you have to handle each specimen, risking more accidental damage.

Most museums use a system of air-tight, steel cabinets and wooden, glass-topped drawers. Into these drawers you can drop small, standard-sized, card boxes lined with foam called unit-trays. Each tray is sized to be a multiple of the smallest unit so that they can be swapped around and organized into neat rows. Each unit-tray then contains 1 species.

You can easily move the boxes around and slot in larger or smaller units as your collection grows without actually handling any pins. Also you reduce the amount of handling each specimen gets in its lifetime because routine checking of a species can be done by just picking out the whole unit tray and holding it under the microscope.

Pros:

it’s easier to quickly see into the glass-topped drawer

reduced individual specimen handling

much easier to add new species

much easier to rearrange your collection

broken body-parts tend to stay in the appropriate tray

the unit-trays give added protection while the specimens are out of the drawer

Cons:

takes up more space

more difficult to carry around and take to workshops etc.

it is much more expensive

Pests

As I mentioned earlier, pests are a worry for all collectors – if museum/carpet beetle (Anthinus sp.) get into a collection they can reduce your prized specimens to dust in a matter of weeks. But if you are worried about pests you can always freeze your specimens in a domestic freezer (2-4 weeks at -20C will do the trick). Museums do this as a matter of course because many insecticide chemicals (naphthalene etc) are now frowned-upon or banned. Just place your box in a black-plastic bag and expel as much air as possible before sealing it up so that air and moisture can’t get in during the freezing process. When you remove them after 2-4 weeks you should allow the bag+box to reach room temperature before opening the bag or box and letting air in – otherwise condensation will form on your specimens!

Posting specimens

Sometimes it might be necessary to send pinned specimens to another expert for identification. This need not be a problem as long as you have good, sturdy boxes with a deep (9mm) later of dense foam attached firmly to the bottom. Into this pin your specimens and if you think the stages might spin you can hold them in place with more 38mm stage-pins.

It is also possible to send specimens from Malaise trapping in glass tubes of alcohol protected by plenty of cotton wool and a sturdy cardboard box. But this might cause some raised eyebrows if you send them internationally because customs never like to see alcohol being brought into the country, whether it has flies in or not!

Conclusion

Whatever you do, start by thinking hard about the groups you are studying and adapt what I have said for your own circumstances. There are no hard and fast rules other than the 2 points I made in the opening paragraphs – protect your specimen and present it so that it can be seen easily. Also, remember that you need to have a system that suits your circumstances and your lifestyle – if it is too much hassle you won’t enjoy your hobby and you won’t have the time to do the really enjoyable and exciting parts of the work. Lastly, if you have any comments you’d like to make or any suggestions for improvement then feel free to leave a comment somewhere and get in touch!